There is currently a need in drug discovery and development and in general biological research for methods and apparatus for accurately performing cell-based assays. Cell-based assays are advantageously employed for assessing the biological activity of chemical compounds.
In addition, there is a need to quickly and inexpensively screen large numbers of chemical compounds. This need has arisen in the pharmaceutical industry where it is common to test chemical compounds for activity against a variety of biochemical targets, for example, receptors, enzymes and nucleic acids. These chemical compounds are collected in large libraries, sometimes exceeding one million distinct compounds. The use of the term chemical compound is intended to be interpreted broadly so as to include, but not be limited to, simple organic and inorganic molecules, proteins, peptides, nucleic acids and oligonucleotides, carbohydrates, lipids, or any chemical structure of biological interest.
In the field of compound screening, cell-based assays are run on populations of cells. The measured response is usually an average over the cell population. For example, a popular instrument used for ion channel assays is disclosed in U.S. Pat. No. 5,355,215. A typical assay consists of measuring the time-dependence of the fluorescence of an ion-sensitive dye, the fluorescence being a measure of the intra-cellular concentration of the ion of interest which changes as a consequence of the addition of a chemical compound. The dye is loaded into the population of cells disposed on the bottom of the well of a multiwell plate at a time prior to the measurement.
In general, the response of the cells is heterogeneous in both magnitude and time. This variability may obscure or prevent the observation of biological activity important to compound screening. Heterogeneity may result from either physiological or genetic differences in cells, or from experimental sources. A method that mitigates, compensates for, or even utilizes the variations would enhance the value of cell-based assays in the characterization of the pharmacological activity of chemical compounds.
Quantification of the response of individual cells circumvents the problems posed by the non-uniformity of that response of a population of cells. Consider the case where a minor fraction of the population responds to the stimulus. A device that measures the average response will have less sensitivity than one determining individual cellular response. However, analysis of the responses of individual cells will be time-consuming in the case of populations of large cell count.
The cell cycle is of key importance to many areas of drug discovery. On the one hand this fundamental process provides the opportunity to discover new targets for anticancer agents and improved chemotherapeutics, but on the other hand drugs and targets in other therapeutic areas must be tested for undesirable effects on the cell cycle. Historically, a wide range of techniques have been developed to study the cell cycle both as a global biochemical process and at the molecular level.
Known methods include those that produce data describing the proliferative activity of a cell population.
Measuring the incorporation of [14C]- or [3H]-thymidine (Regan, J. D. and Chu, E. H. (1966) “A convenient method for assay of DNA synthesis in synchronized human cell cultures” J. Cell Biol. 28, 139-143) by scintillation counting was one of the earliest methods of determining cell proliferation, and is still widely used today. More recent developments (Graves, R. et al. (1997) “Noninvasive, real-time method for the examination of thymidine uptake events—application of the method to V-79 cell synchrony studies” Anal. Biochem. 248, 251-257) have allowed thymidine incorporation to be measured in a homogeneous microplate assay format.
Several non-radioactive alternatives to thymidine incorporation assays have been developed. These include enzyme-linked immunosorbent assay (ELISA) nucleotide bromo-deoxyunridine (BrdU) (Perros, P. and Weightman, D. R. (1991) “Measurement of cell proliferation by enzyme-linked immunosorbent assay (ELISA) using a monoclonal antibody to bromodeoxyuridine. Cell. Prolif. 24, 517-523; Wemme, H. et al. (1992) “Measurement of lymphocyte proliferation: critical analysis of radioactive and photometric methods” Immunobiology 185, 78-89) into replicating DNA, and staining of proliferation-specific antigens such as Ki-67 (Frahm, S. O. et al (1998) “Improved ELISA proliferation assay (EPA) for the detection of in vitro cell proliferation by a new Ki-67-antigen directed monoclonal antibody (Ki-S3)” J. Immunol. Methods 211, 43-50).
Colourimetric methods based on substrate conversion (Mosmann, T. (1983) “Rapid colourimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays” J. Immunol. Methods 65, 55-63; Roehm, N. W. et al. (1991) “An improved colourimetric assay for cell proliferation and viability utilizing the tetrazolium sal XTT” J. Immunol. Methods 142, 257-265) by mitochondrial and other cellular enzymes are also used to measure cell growth. Although these assays are often referred to as cell-proliferation assays, strictly speaking they are cell-mass assays. Unlike measuring thymidine or BrdU incorporation, these assays do not provide any inherent measure of cell cycle progression, and give only a measure of cell mass ie. increase in cell number, relative to another population.
Other methods for measuring cell proliferation (i.e. increasing cell numbers) have been reported based on measuring electrical impedance (Upadhyay, P. and Bhaskar, S. (2000) “Real time monitoring of lymphocyte proliferation by an impedance method” J. Immunol. Methods 244, 133-137), dissolved oxygen (Wodnicka, M. et al (2000) “Novel fluorescent technology platform for high throughput cytotoxicity and proliferation assays” J. Biomol. Screen. 5, 141-152) and others. However, as for the colourimetric assays discussed above, these do not directly report cell cycle parameters and have not been widely adopted.
All of the above methods provide data on the overall proliferation within a cell population under examination, but do not identify the status of individual cells. Adaptation of these assays to imaging, for example by micro-autoradiography of [3H]- or [14C]-thymidine incorporation (Dormer, P. (1981) “Quantitative carbon-14 autoradiography at the cellular level: principles and application for cell kinetic studies” Histochem. J. 13, 161-171) or by immunocytochemical or immunofluorescence detection of BrdU (Dolbeare, F. (1995) “Bromodeoxyuridine: a diagnostic tool in biology and medicine, Part I: historical perspectives, histochemical methods and cell kinetics” Histochem. J. 27, 339-369) permits identification of cells that have traversed S phase, but does not yield information on the cell cycle position of other cells under analysis.
To determine the cell cycle status of all cells in a population it is a prerequisite that the analytical technique can resolve at least to the level of a single cell. Of the two qualifying techniques available, flow cytometry and microscopy, flow cytometry has become firmly established as the standard method for analysing cell cycle distribution.
The DNA content of cell nuclei varies through the cell cycle in a predictable fashion—cells in G2 or M have twice the DNA content of cells in G1, and cells undergoing DNA synthesis in S phase have an intermediate amount of DNA. Consequently, staining of cellular DNA with propidium iodide (Nairn, R. C. and Rolland, J. M. (1980) “Fluorescent probes to detect lymphocyte activation” Clin. Exp. Immunol. 39, 1-13) or other fluorescent dyes (Smith, P. J. et al (2000) “Characteristics of a novel deep red/infrared fluorescent cell-permeant DNA probe, DRAQ5, in intact human cells analyzed by flow cytometry, confocal and multiphoton microscopy” Cytometry 40, 280-291) that are compatible with live cells, followed by flow cytometry permits measurement of the relative proportion of cells in G1, S and G2/M phases. However, analysis by propidium iodide staining and flow cytometry is necessarily destructive and hence requires multiple samples to study cell cycle progression, which can become rate limiting where many hundreds of samples are to be analysed. In addition, flow cytometry does not yield fine resolution of cell cycle position in G2/M as the DNA content is the same in all cells.
A combination of DNA staining with pulsed BrdU incorporation can be used to resolve the cell cycle position further (Dolbeare, F. et al. (1983) “Flow cytometric measurement of total DNA content and incorporated bromodeoxyuridine” Proc. Natl. Acad. Sci. U.S.A. 80, 5573-5577). Dual-parameter analysis of DNA staining and/or BrdU incorporation can also be used with antibodies to cell-surface markers to profile cell cycle distribution in a defined subpopulation of cells (Mehta, B. A. and Maino, V. C. (1997) “Simultaneous detection of DNA synthesis and cytokine production in staphylococcal enterotoxin B activated CD4+T lymphocytes by flow cytometry” J. Immunol. Methods 208, 49-59; see also Johannisson, A. et al. (1995) “Activation markers and cell proliferation as indicators of toxicity: a flow cytometric approach” Cell Biol. Toxicol. 11, 355-366; see also Penit, C. and Vasseur, F. (1993) “Phenotype analysis of cycling and postcycling thymocytes: evaluation of detection methods for BrdUrd and surface proteins” Cytometry 14, 757-763).
Although to date flow cytometry has remained the dominant method for analysing the cell cycle, many of the above techniques have also been applied to microscopic analyses (Gorczyca, W. et al. (1996) “Laser scanning cytometer (LSC) analysis of fraction of labeled mitoses (FLM)” Cell Prolif. 29, 539-547; Clatch, R. J. and Foreman, J. R. (1998) “Five-colour immunophenotyping plus DNA content, analysis by laser scanning cytometry” Cytometry 34, 36-38).
The techniques described above all provide information in various forms from a single point in time (e.g. propidium iodide staining for DNA content) or integrated over a period of time (e.g. thymidine or BrdU incorporation). One further technique, cell-division tracking (Nordon, R. E. et al. (1999) “Analysis of growth kinetics by division tracking” Immunol. Cell Biol. 77, 523-529; Lyons, A. B. (1999) “Divided we stand: tracking cell proliferation with carboxyfluorescein diacetate succinimidyl ester” Immunol. Cell. Biol. 77, 509-515), allows the replicative history of a cell population to be analysed. In this method cells are loaded with a fluorescent dye such as carboxy-fluorescein diacetate succinimidyl ester (CFSE), which is partitioned between daughter cells at each successive round of cell division with a twofold reduction in fluorescence. Subsequent analysis of cell fluorescence by flow cytometry reveals the number of cell divisions undergone by each cell in the population. This technique has also been used in multi-parameter analyses combined with BrdU and proliferation-marker staining (Hasbold, J. and Hodgkin, P. D. (2000) “Flow cytometric cell division tracking using nuclei” Cytometry 40, 230-237).
International patent application WO 01/11341 describes a method for the automated measurement of the mitotic index of cells using fluorescence imaging. The technique involves immunoflourescence which reports specifically on mitotic cells by signals emitted from the cell nuclei, dependent upon the phosphorylation of histone H3. A mitotic index is determined by detecting the number of mitotic cells compared with the number of nuclei detected in a separate fluorescence channel. The technique involves simply counting cells having a signal above a given threshold, and is unsuited for the detection of cell cycle phases other than mitosis. Furthermore, the signal thresholds have to be predetermined, or entered by an operator.
The application of GFP and imaging techniques to cell cycle analysis has enabled significant advances to be made in understanding the timing of the molecular events that control the cell cycle. Fusing. GFP with key cell-cycle-control proteins has provided significant insights into the molecular organisation behind the cell cycle (see (Raff, J. W. et al (2002) “The roles of Fzy/Cdc20 and Fzr/Cdh1 in regulating the destruction of cyclin B in space and time” J. Cell Biol. 157, 1139-1149; Zeng, Y. et al. (2000) “Minimal requirements for the nuclear localization of p27(Kip1), a cyclin-dependent kinase inhibitor” Biochem. Biophys. Res. Commun. 274, 37-42; Huang, J. and Raff. J. W. (1999) “The disappearance of cyclin B at the end of mitosis is regulated spatially in Drosophila cells” EMBO J. 18, 2184-2195; Weingartner, M. et al. (2001) “Dynamic recruitment of Cdc2 to specific microtubule structures during mitosis” Plant Cell 13, 1929-1943; Arnaud, L. et al. (1998) “GFP tagging reveals human Polo-like kinase 1 at the kinetochore/centromere region of mitotic chromosomes” Chromosoma 107, 424-429) and other cellular components (Kanda, T. et al. (1998) “Histone-GFP fusion protein enables sensitive analysis of chromosome dynamics in living mammalian cells” Curr. Biol. 8, 377-385; Reits, E. A. et al. (1997) “Dynamics of proteasome distribution in living cells” EMBO J. 16, 6087-6094; Tatebe, H. et al. (2001) “Fission yeast living mitosis represented by GFP-tagged gene products” Micron 32, 67-74)). However, although these specialised approaches provide valuable data on the mechanisms and components involved, they are not generic methods for monitoring the cell cycle.
Another purpose of cell cycle analysis (and for example cyclin cell lines) is to first classify the cells in the population, then to perform analysis of other parameters on each subpopulation separately using reporters in other channels. Cells at different stages will respond differently to different compounds (e.g. cell surface receptors cannot be activated in mitotic cells.)